Method for culturing skeletal muscle for tissue engineering

ABSTRACT

The invention provides a nutrient medium composition and associated methods for lengthening the useful life of a culture of muscle cells. Disclosed is a method of culturing mammalian muscle cells, including preparing one or more carriers coated with a covalently bonded monolayer of trimethoxy-silylpropyl-diethylenetriamine (DETA); verifying DETA monolayer formation by one or more associated optical parameters; suspending isolated fetal rat skeletal muscle cells in serum-free medium according to medium composition 1; plating the suspended cells onto the prepared carriers at a predetermined density; leaving the carriers undisturbed for cells to adhere to the DETA monolayer; covering the carriers with a mixture of medium 1 and medium 2; and incubating. A cell nutrient medium composition includes Neurobasal, an antibiotic-antimycotic composition, cholesterol, human TNF-alpha, PDGF BB, vasoactive intestinal peptides, insulin-like growth factor 1, NAP, r-Apolipoprotein E2, purified mouse Laminin, beta amyloid, human tenascin-C protein, rr-Sonic hedgehog Shh N-terminal, and rr-Agrin C terminal.

RELATED APPLICATION

This application claims priority from co-pending provisional application Ser. No. 61/171,968, which was filed on 23 Apr. 2009, and which is incorporated herein by reference in its entirety.

STATEMENT OF GOVERNMENT RIGHTS

Development of the present invention was supported, at least in part, by a grant from the U.S. Government. Accordingly, the Government may have certain rights in the invention, as specified by law.

FIELD OF THE INVENTION

The present invention relates to the field of tissue engineering and, more particularly, to a system and method of extending the in vitro useful life of a culture of muscle cells.

BACKGROUND OF THE INVENTION

Skeletal muscle differentiation and maturation is a complex process involving the synergy of different growth factors and hormones interacting over a broad time period [1-11]. The differentiation process is further complicated by neuronal innervation, where neuron to muscle cell signaling can regulate myosin heavy chain (MHC) gene expression and acetylcholine receptor clustering [12-18]. Consequently, understanding of the role of the growth factors, hormones and cellular interactions in skeletal muscle differentiation would be a key step in generating physiologically relevant tissue engineering constructs, developing advanced strategies for regenerative medicine and integrating functional skeletal muscle with bio-hybrid MEMS devices for non-invasive interrogation in high-throughput screening technologies.

In order for skeletal muscle myotubes developed in vitro to be useful in tissue engineering applications, they must exhibit as many of the functional characteristics of in vivo skeletal muscle fibers as possible. During muscle fiber development in vivo, several critical structural changes occur that indicate functional maturation of the extrafusal myotubes. These changes include sarcomere organization, clustering and colocalization of ryanodine (RyR) and dihydropyridine (DHPR) receptors and MHC class switching [19-23]. Each of these structural changes reflects the physiological maturation of the skeletal muscle and is critical for consistent muscular contraction. For example, organization of the contractile proteins myosin and actin into sarcomeric units gives skeletal muscle myotubes organized and structured contraction, a property lacking in smooth muscle. The organization of sarcomeres in skeletal muscle gives rise to anisotropic and isotropic bands of proteins (A and I bands) and gives skeletal muscle a striated appearance. The clustering and colocalization of RyR and DHPR is indicative of transverse tubule (T-tubule) biogenesis and excitation contraction coupling. This developmental step structurally links electrical excitation to the internal contractile system by providing close apposition of DHPR located in the T-tubule and RyR located in the sarcoplasmic reticulum. Finally, a properly functioning skeletal muscle must express the appropriate MHC proteins required for the task it must perform. For example, different muscle fibers express different MHC proteins depending on the rate of contraction and force generation required by the work to be done. Consequently, skeletal muscle fibers change their MHC expression profiles to best meet the requirements of the body as it matures. Without these modifications, an in vitro model of skeletal muscle maturation cannot achieve full physiological relevance.

One approach for identifying the role of specific growth factors and hormones in muscle differentiation is to develop an in vitro model system consisting of a serum-free medium supplemented with the factors of interest. Such a model provides the opportunity to evaluate the role of each factor individually or in combination with others known or believed to be important in skeletal muscle development. For example, the concentration and/or temporal application of medium components in order to influence the maturation of extrafusal fiber or intrafusal fiber subtypes could be easily investigated.

Employing a non-biological growth substrate such as trimethoxy-silylpropyl-diethylenetriamine (DETA) provides an additional measure of control. DETA is a silane molecule that forms a covalently bonded monolayer on glass coverslips, resulting in a uniform, non-hydrophilic surface for cell growth. The use of DETA surfaces is advantageous from a tissue engineering perspective because it can be covalently linked to virtually any hydroxolated surface, it is amenable to patterning using standard photolithography and it promotes long-term cell survival because it is non-digestible by matrix metalloproteinases secreted by the cells [24, 25].

Previously, studies have demonstrated the usefulness of the DETA silane substrate for in vitro culture systems. Interesting features of the DETA silane are that its molecular geometry does not allow for an ordered nanolayer and may partially mimic the three dimensional features of an extracellular matrix, which may be responsible for robust growth of different cell types on this synthetic substrate [24-31]. Additionally, DETA's non-biological nature supports the analysis of ECM proteins secreted by the cell in response to different in vitro conditions.

We earlier developed a defined system that promoted differentiation of different skeletal muscle phenotypes and resulted in the formation of contractile myotubes. This resulted in short-term survival of the myotubes [25, 28]. We also have developed a novel bio-hybrid technology to integrate functional myotubes with cantilever based bio-MEMS devices for the study of muscle physiology, neuromuscular junction formation and bio-robotics applications for use in a model of the stretch reflex arc [32]. More recently, using our defined model system, we have achieved a significant breakthrough by creating mechanosensitive intrafusal myotubes in vitro [33]. The intrafusal fibers differentiated upon addition of neuregulin 1-□-1 to serum-free medium in our defined system. Intrafusal fibers are the myotubes present in the muscle spindle which functions as the sensory receptor of the stretch reflex circuit [16] and combined with extrafusile fibers represent the primary component necessary to reproduce functional muscle function in vitro.

This system has been utilized as a model for different developmental and functional applications, however, further improvements are necessary to enhance the physiological relevance of the skeletal muscle myotubes [32, 33]. Specifically, in order to create a working model of the stretch reflex arc, myotubes are needed that more accurately represent extrafusal fibers in vivo. A more advanced developmental system for skeletal muscle would have applications in basic science research and tissue engineering. In this study, we have demonstrated sarcomere assembly, the development of the excitation-contraction coupling apparatus and myosin heavy chain (MHC) class switching.

The results disclosed herein suggest we have discovered a group of biomolecules that act together as a molecular switch promoting the transition from embryonic to neonatal MHC expression as well as other structural adaptations resulting in the maturation of skeletal muscle in vitro. The discovery of these biomolecular switches will be a powerful tool in regenerative medicine and tissue engineering applications such as skeletal muscle tissue grafts. It should also be useful in higher content high-throughput screening technology.

SUMMARY OF THE INVENTION

With the foregoing in mind, the present invention advantageously provides a method of culturing mammalian muscle cells. The method of the invention includes preparing one or more carriers coated with a covalently bonded monolayer of trimethoxy-silylpropyl-diethylenetriamine (DETA). This is followed by verifying DETA monolayer formation by one or more associated optical parameters. The method continues by suspending isolated fetal rat skeletal muscle cells in serum-free medium according to medium composition 1, as set forth below in further detail, then plating the suspended cells onto the prepared carriers at a predetermined density. The method then calls for leaving the carriers undisturbed while the plated cells adhere to the DETA monolayer and covering the carriers with a mixture of medium composition 1 and medium composition 2, both as described below. Finally, the method ends by incubating the carriers.

In the method, the one or more carriers typically comprise glass cover slips. Those of skill in the art should understand that verifying is accomplished by an optical contact angle goniometer in the present invention, which may also include verifying by X-ray photoelectron spectroscopy (XPS). In the method, verifying may be accomplished by both an optical contact angle goniometer and by XPS.

Our current cantilever system is designed for force measurements of contracting muscle cells and uses laser optics as a readout system [136]. Alternatively, piezoresistive and piezoelectric approaches are the most widely applied techniques for measuring stress applied on microcantilevers [137] and could be easily adapted to the present invention by those of ordinary skill in the art. The advantage is that the mechanical device and the read our electronics can be implemented in the same integrated circuit. Replacing the optical readout with piezoelements will reduce the size and complexity of our current cantilever system.

Those skilled in the art will know that piezoelectricity is the ability of certain materials (crystals and certain ceramics) to generate an electric potential in response to applied mechanical stress [132]. The piezoelectric effect is used in various sensors to measure stresses or geometrical deformations in various mechanical devices. The reverse piezoelectric effect turns piezoelectric materials into actuators, when an external voltage is applied to the crystal [133]. Piezoelectric materials are e.g. quartz, bone, sodium tungstate, zinc oxide, or lead zirconate titanate (PZT) [134]. A similar effect is the piezoresistive phenomenon. When subjected to mechanical stress, these materials change their resistivity [135].

Culture aspects of the method include wherein plating the muscle cells is at a density of approximately from 700 to 1000 cells/mm2. Then, leaving the carriers undisturbed continues for approximately one hour and incubating is effected under physiologic conditions and may best be accomplished at approximately 37° C. in an air atmosphere with about 5% CO₂ and 85% humidity. The culture is then covered with a mixture of approximately equal volumes of medium composition 1 and medium composition 2. Preferably, an initial complete change of the medium covering the carriers is accomplished by substituting NBactiv4 medium during incubation. Moreover, after the initial complete change of medium, changing every three days more than half of the medium covering the carriers is preferred and it is most preferred changing every three days approximately three quarters of the medium covering the carriers.

Another embodiment of the present invention includes a method of culturing mammalian muscle cells which comprises allowing mammalian fetal muscle cells suspended in medium according to composition 1to adhere to a monolayer of covalently bonded DETA formed on an underlying carrier surface and incubating the adhered cells covered in a mixture of approximately equal volumes of medium composition 1 and medium composition 2.

In the methods of the invention, the mammalian fetal muscle cells may comprise fetal rat cells and the underlying carrier surface may comprise a glass cover slip. Incubating is preferably under physiological conditions, typically at approximately 37° C. in an atmosphere of air with about 5% CO₂ and 85% humidity.

In the alternate embodiment of the invention, the method includes changing the covering medium to NBactiv4, preferably after approximately four days of incubation. Thereafter, the method calls for changing every three days more than half of the medium covering the carriers and preferably about three quarters of the medium covering the carriers.

Also part of the invention is a new cell culture medium composition which includes NBactiv4, an antibiotic-antimycotic composition, cholesterol, human TNF-alpha, PDGF BB, vasoactive intestinal peptides, insulin-like growth factor 1, NAP, r-Apolipoprotein E2, purified mouse Laminin, beta amyloid, human tenascin-C protein, rr-Sonic hedgehog Shh N-terminal, and rr-Agrin C terminal. This medium composition may be amplified with G5 supplement, VEGF, acidic fibroblast growth factor, heparin sulphate, LIF, rat plasma Vitronectin, CNTF, GNDF, NT-3, NT-4, BDNF and CT-1.

BRIEF DESCRIPTION OF THE DRAWINGS

Some of the features, advantages, and benefits of the present invention having been stated, others will become apparent as the description proceeds when taken in conjunction with the accompanying drawings, presented for solely for exemplary purposes and not with intent to limit the invention thereto, and in which:

FIG. 1. is a schematic diagram of a culture protocol according to an embodiment of the present invention;

FIG. 2 A, B, C and D provide phase pictures of 50 day old myotubes in culture; red arrows showing characteristic striations in most of the myotubes; scale bar equals 75 μm;

FIG. 3 shows myotubes stained with antibodies against embryonic myosin heavy chain (F 1.652) proteins at day 50; scale bar is 75 μm; A) panel showing phase+fluorescence picture of the myotubes; B) another view of panel A but observed only under fluorescence; white arrows showing the striations; C) panel showing image of myotubes under phase+fluorescence illumination; D) shows panel C observed only under fluorescence illumination; E) panel showing phase+fluorescence picture of the myotubes (white arrow indicating the striations); F) panel E observed only in fluorescence light (white arrow indicating the striations); G) panel showing phase+fluorescence picture of the myotubes (white arrow indicating the striations); H) panel G observed only under fluorescence light (white arrow indicating the striations);

FIG. 4 shows myotubes immunostained with neonatal myosin heavy chain (N3.36) and alpha-bungarotoxin at day 50; scale bar is 75 μm; A) is a phase picture of 2 myotubes indicated by white arrows; B) both the myotubes shown in phase (panel A) have acetylcholine receptor clustering indicated by alpha-bungarotoxin staining; C) only one myotube out of the two seen in panel A stained for N3.36; D) double stained image of panel A with alpha-bungarotoxin and N3.36; E) phase image of 6 myotubes indicated by white arrows; F) all the myotubes shown in phase (panel E) have acetylcholine receptor clustering shown by alpha-bungarotoxin staining; G) none of the myotubes in panel E stained for N3.36; H) I) and J) show differential staining of the myotubes with N3.36; K) L) and M) showing differential staining of the myotubes with N3.36;

FIG. 5 illustrates ryanodine receptor and DHPR receptor clustering in 30 days old skeletal muscle culture (scale bar 75 μm); A) phase and fluorescent-labeled picture of the myotubes; B) merged fluorescence picture of the ryanodine receptor (green) and DHPR receptor (red) clustering on the myotubes shown in panel A; C) ryanodine receptor (green) on the myotubes shown in panel A; D) DHPR receptors on the myotubes shown in panel A; E) phase and fluorescent-labeled picture of the myotubes; F) merged fluorescent picture of the ryanodine receptor (green) and DHPR receptor (red) clustering on the myotubes (panel E); G) ryanodine receptor (green) on the myotubes (panel E); H) DHPR receptors on the myotubes (panel E); I) phase and fluorescent-labeled picture of the myotubes; J) K) and L) show merged fluorescence pictures of the ryanodine receptor (green) and DHPR receptor (red) clustering on the myotubes (panel I) at three different planes (white arrows indicate the striations and the receptor clustering);

FIG. 6 shows ryanodine receptor and DHPR receptor clustering in 100 days old skeletal muscle culture (scale bar: 75 μm); A) shows phase and fluorescent-labeled picture of the myotubes; B) is a merged fluorescence picture of the ryanodine receptor (green) and DHPR receptor (Red) clustering on the myotubes (panel A); C) shows ryanodine receptor (green) on the myotubes (panel A); D) shows DHPR receptors on the myotubes (panel A); E and F show views of the same panels at different planes showing the merged fluorescent picture of the ryanodine receptor (green) and DHPR receptor (red) clustering on the myotubes;

FIG. 7 depicts patch clamp electrophysiology of the myotubes, wherein A shows representative voltage clamp trace obtained after patching a 48 days old myotube in culture; B shows representative current clamp trace of the same myotube for which voltage clamp trace had been obtained (inset showing the picture of patched myotubes).

DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENT

The present invention will now be described more fully hereinafter with reference to the accompanying drawings, in which preferred embodiments of the invention are shown. Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art to which this invention pertains. Although methods and materials similar or equivalent to those described herein can be used in the practice or testing of the present invention, suitable methods and materials are described below. Any publications, patent applications, patents, or other references mentioned herein are incorporated by reference in their entirety. In case of conflict, the present specification, including any definitions, will control. In addition, the materials, methods and examples given are illustrative in nature only and not intended to be limiting. Accordingly, this invention may, however, be embodied in many different forms and should not be construed as limited to the illustrated embodiments set forth herein. Rather, these illustrated embodiments are provided so that this disclosure will be thorough and complete, and will fully convey the scope of the invention to those skilled in the art. Other features and advantages of the invention will be apparent from the following detailed description, and from the claims.

Materials and Methods Surface Modification and Characterization

Glass coverslips (Thomas Scientific 6661F52, 22×22 mm No. 1) were cleaned using an O₂ plasma cleaner (Harrick PDC-32G) for 20 minutes at 100 mTorr. The DETA (United Chemical Technologies Inc. T2910KG) films were formed by the reaction of the cleaned glass surface with a 0.1% (v/v) mixture of the organosilane in freshly distilled toluene (Fisher T2904). The DETA coated coverslips were then heated to approximately QQ 100° C., rinsed with toluene, reheated to approximately 100° C., and then oven dried [28]. Surfaces were characterized by contact angle measurements using an optical contact angle goniometer (KSV Instruments, Cam 200) and by X-ray photoelectron spectroscopy (XPS) (Kratos Axis 165). XPS survey scans, as well as high-resolution N1s and C1s scans utilizing monochromatic Al Kα excitation were obtained [28].

Skeletal Muscle Culture and Serum Free Medium

The skeletal muscle was dissected from the thighs of the hind limbs of fetal rats (17-18 days old). The tissue was collected in a sterile 15 mL centrifuge tube containing 1 mL of phosphate-buffered saline (calcium- and magnesium-free) (Gibco 14200075). The tissue was enzymatically dissociated using 2 mL of 0.05% of trypsin-EDTA (Gibco 25300054) solution for 30 minutes in a 37□ C water bath at 50 rpm. After 30 minutes the trypsin solution was removed and 4 mL of Hibernate E+10% fetal bovine serum (Gibco 16000044) was added to terminate the trypsin reaction. The tissue was then mechanically triturated with the supernatant being transferred to a 15 mL centrifuge tube. The same process was repeated two times by adding 2 mL of L15+10% FBS each time. The 6 mL cell suspension obtained after mechanical trituration was suspended on a 2 mL, 4% BSA (Sigma A3059) (prepared in L15 medium) cushion and centrifuged at 300 g for 10 minutes at 4° C. The pellet obtained was washed 5 times with L15 medium then resuspended in 10 mL of L15 and plated in 100 mm uncoated dishes for 30 minutes. The non-attached cells were removed and then centrifuged on a 4% BSA cushion [28]. The pellet was resuspended in serum-free medium according to the protocol illustrated in FIG. 1 and plated on the coverslips at a density of 700-1000 cells/mm². The serum-free medium containing different growth factors and hormones was added to the culture dish after one hour. The final medium was prepared by mixing medium one (Table 1) and medium two (Table 2) in a 1:1 v/v ratio. FIG. 1 indicates the flowchart of the culture protocol. Tables 1 and 2 indicated the growth factor and hormone supplement compositions of medium one and medium two. The cells were maintained in a 5% CO₂ incubator (relative humidity 85%). The full medium was replaced after four days with NBactiv4 medium according to the protocol in FIG. 1 [34]. Thereafter three-fourth of the medium was changed every three days with NBactiv4.

Immunocytochemistry of Skeletal Muscle Myotubes

Coverslips were prepared for immunocytochemical analysis as previously described. Briefly, coverslips were rinsed with PBS, fixed in −20□ C methanol for 5-7 min, washed in PBS, incubated in PBS supplemented with 1% BSA and 0.05% saponin (permeabilization solution) for 10 minutes, and blocked for 2 h with 10% goat serum and 1% BSA. Cells were incubated overnight with primary antibodies against embryonic myosin heavy chain (F1.652) (dilution>1:5), neonatal myosin heavy chain (N3.36) (1:5) (Developmental Studies Hybridoma Bank), ryanodine receptor (AB9078, Millipore) (1:500) and dihydropyridine binding complex (α1-Subunit) (MAB 4270, Millipore) (1:500) diluted in the blocking solution. Cells were washed with PBS and incubated with the appropriate secondary antibodies for two hours in PBS. After two hours the coverslips were rinsed with PBS and mounted on glass slides and evaluated using confocal microscopy [25, 28, 31].

AChR Labeling of Myotubes

AChRs were labeled as described previously by incubating cultures with 5×10-8 M of α-bungarotoxin, Alexa Fluor® 488 conjugate (B-13422; Invitrogen) for 1.5 h at 37° C. [12, 31]. Following incubation in α-bungarotoxin, the cultures were fixed as above for subsequent staining with embryonic myosin heavy chain (F1.652) antibodies.

Patch Clamp Electrophysiology of the Myotubes

Whole-cell patch clamp recordings were performed in a recording chamber located on the stage of a Zeiss Axioscope 2FS Plus upright microscope as described previously [25, 33]. The chamber was continuously perfused (2 ml/min) with the extracellular solution (Leibovitz medium, 35° C.). Patch pipettes were prepared from borosilicate glass (BF150-86-10; Sutter, Novato, Calif.) with a Sutter P97 pipette puller and filled with intracellular solution (K-gluconate 140 mM, EGTA 1 mM, MgCl₂ 2 mM, Na₂ATP 2 mM, phosphocreatine 5 mM, phosphocreatine kinase 2.4 mM, Hepes 10 mM; pH=7.2). The resistance of the electrodes was 6-8MΩ. Voltage clamp and current clamp experiments were performed with a Multiclamp 700A amplifier (Axon Laboratories, Union City, Calif.). Signals were filtered at 2 kHz and digitized at 20 kHz with an Axon Digidata 1322A interface. Data recording and analysis were done with pClamp 8 software (Axon Laboratories). Membrane potentials were corrected by subtraction of a 15 mV tip potential, which was calculated using Axon's pClamp 8 program. Sodium and potassium currents were measured in voltage clamp mode using voltage steps from a −85 mV holding potential. Action potentials were evoked with 1 second depolarizing current injections from a -85 mV holding potential [25, 28].

Results DETA Surface Modification and Characterization

Static contact angle and XPS analysis was used for the validation of the surface modifications and for monitoring the quality of the surfaces. Stable contact angles (40.64±2.9/mean±SD) throughout the study indicated high reproducibility and quality of the DETA surfaces and were similar to previously published results [24, 25, 28, 29, 31]. Based on the ratio of the N (401 and 399 eV) and the Si 2p3/2 peaks, XPS measurements indicated that a reaction-site limited monolayer of DETA was formed on the coverslips [35].

Development of the Serum Free Medium Formulation and Culture Timeline for Long-Term Survival and Maturation of Myotubes

The serum free medium composition was developed empirically. The final medium is derived from two different medium compositions described in Tables 1 and 2. Table 1 constitutes the same medium composition used previously for a motoneuron-muscle co-culture and adult spinal cord neurons culture [26, 27, 30, 31]. Table 2 is composed of twelve additional factors that had been shown to promote skeletal muscle maturation and neuromuscular junction formation separately. The final medium was prepared by mixing these two media in a 1:1 v/v ratio. After first 4 days of culture the whole medium was replaced with NBactiv4 medium [34]. Thereafter, every three days three-fourth medium was changed with NBactiv4. The culture technique has been illustrated in the flowchart (FIG. 1).

Using this new medium formulation and timeline, myotubes were successfully cultured for more than 50 days. FIG. 2 indicates 50 days old myotubes in culture. As the myotubes aged and grew they began to form the characteristic anisotropic (A band) and isotropic (I band) banding pattern observed with in vivo muscle fibers [22, 23]. This banding pattern is caused by differential light diffraction due to the organization of myofibril proteins forming sarcomeres within the myotubes [22, 23]. The arrowheads in the images (FIG. 2A-D) indicate myotubes where sarcomeric organization has occurred and is visualized by the appearance of A and I bands.

Myotube Expression of Fetal Myosin Heavy Chain

The myotubes formed were evaluated for the expression of fetal MHC to establish a baseline as comparison to our previous results [28]. In FIG. 3, the myotubes phenotypes formed at approximately day 50 in vitro are shown. The myotubes ranged from having clustered nuclei (FIG. 3A-D) to having diffuse nuclear organization (FIG. 3E-H). The arrowheads in the images indicate the characteristic striations.

Differential Expression of Neonatal MHC Protein in the Myotubes

In order to determine if the myotubes were maturing in a physiologically relevant way as they aged in vitro, the expression of neonatal MHC protein was evaluated. After approximately 50 days in vitro 25% of the myotubes expressed neonatal MHC (FIG. 4A-M). Additionally, the myotubes were stained for clustering of acetylcholine receptors (AChR) using alpha bungarotoxin (FIG. 5B,F). This clustering of the AChR receptors, induced by the motoneuron protein agrin in vivo, are locations on the myotube where neuromuscular junction formation occurs.

Formation of the Excitation—Contraction Coupling Apparatus

The presence of ryanodine (RyR) receptor and dihydropyridine (DHPR) receptor clusters, as well as their colocalization in vivo, represents the development of excitation-contraction coupling apparatus in skeletal muscle myotubes [19, 21-23]. The clustering of both RyR and DHPR receptors was observed on the myotubes after 30 days in culture (FIG. 5A-D). The clustering and colocalization of the RyR+DHPR clusters was observed with different myotube morphologies (FIG. 5E-L). This functional adaptation illustrated that the medium formulation facilitated not only the structural maturation but also the functional maturation of myotubes in this in vitro system. The clustering of the RyR+DHPR receptors was also observed in the 70 day old myotubes, indicating that the older myotubes maintained their functional integrity (FIG. 6A-F).

Myotube Electrophysiology

The myotubes contracted spontaneously in the culture and the contractions began generally by day four and continued throughout the life of the culture. Most of the myotubes expressed functional voltage gated sodium, potassium and calcium ion channels as reported previously [28]. The voltage clamp electrophysiology of the myotubes indicated the inward and outward currents that demonstrate functional sodium and potassium channels (FIG. 7A). The current clamp study indicated the single action potential fired by the myotubes (FIG. 7B).

Discussion

Herein we have documented the development of a system for long-term in vitro functional, skeletal muscle culture. This system was developed in response to a need for more physiologically relevant skeletal muscle myotubes for functional in vitro systems. For our specific research, they were needed for a realistic model of the stretch reflex arc development and to be integrated with bio-MEMS cantilevers for screening applications. The results indicate we achieved three significant structural modifications within the myotubes, causing both the developmental profile and functionality of the fibers to better mimic in vivo physiology. It is believed that this skeletal muscle maturation resulted from modifications to the cell culture technique, a new medium formulation and the use of NBactiv4 as the maintenance medium.

The presently described serum-free medium supplemented with growth factors was developed to support the survival, proliferation and fusion of fetal rat myoblasts into contractile myotubes. The rationale for selecting the growth factors was based on the distribution of their cognate receptors in the developing myotubes in rat fetus [1-11]. Tables 1 and 2 reference the literature where these individual growth factors, hormones and neurotransmitters were observed to support muscle and neuromuscular junction development. The composition in Table 1 is the formulation used for a previously published medium used for motoneuron-muscle co-culture and adult spinal cord neuron culture [26, 27, 30, 31]. Table 2 lists the twelve additional factors we have identified in muscle development and neuromuscular junction formation. The use of NBactiv4 for the maintenance of the cells provided unexpected results in that it significantly improved the survival of the skeletal muscle derived myotubes despite the original development of NBactiv4 for the long-term maintenance and synaptic connectivity of fetal hippocampal neurons in vitro [34].

We observed a ratio of 25% neonatal to 75% embryonic MHC expression of the myotubes, which contrasts with the previous study in which MHC expression was strictly embryonic. We believe that the myotubes matured in this culture system because the long-term survival provided adequate time for the myotubes to respond to the additional growth factors, which activated the necessary signaling pathways to achieve MHC class switching [20]. This suggests that a different growth factor profile could be utilized to activate alternative signaling pathways and drive myotube differentiation down other pathways. For example, the effects of adding steroid hormones like testosterone to the system could be critically examined.

The colocalization of RyR and DHPR clusters in the myotubes indicated the formation of excitation-contraction coupling apparatus and was another indicator of functional maturation in the fibers. Excitation-contraction coupling is the signaling process in muscle by which membrane depolarization causes a rapid elevation of the cytosolic Ca²⁺ generating contractile force [36]. The close proximity of the DHPR and RyR complexes occurs at specialized junctions established between the transverse tubule and sarcoplasmic reticulum (SR) membranes in skeletal muscle myotubes [37]. At these junctions, T-tubule depolarization is coupled to Ca²⁺ release from the SR resulting in muscle contraction [38-40]. This structural adaptation represents a significant functional change due to the fact that excitation-contraction coupling is required for successful extrafusal muscle fiber development as well as neuromuscular junction formation [19, 21-23]. This improved model provides the potential to study excitation-contraction coupling in a defined system as well as myotonic and myasthenic diseases.

Conclusion

The development of sarcomeric structures, the excitation-contraction coupling apparatus and MHC class switching in the skeletal muscle myotubes is a result of the improvements to the model system documented in this research. This improved system along with the new findings support the goal of creating physiologically relevant tissue engineered muscle constructs and puts within reach the goal of functional skeletal muscle grafts. Furthermore, we believe this serum-free culture system will be a powerful tool in developing advanced strategies for regenerative medicine in muscular dystrophies, stretch reflex arc development and integrating skeletal muscle with bio-hybrid prosthetic devices.

Accordingly, in the drawings and specification there have been disclosed typical preferred embodiments of the invention and although specific terms may have been employed, the terms are used in a descriptive sense only and not for purposes of limitation. The invention has been described in considerable detail with specific reference to these illustrated embodiments. It will be apparent, however, that various modifications and changes can be made within the spirit and scope of the invention as described in the foregoing specification and as defined in the appended claims.

REFERENCES

-   1. Arnold H H, Winter B. Muscle differentiation: more complexity to     the network of myogenic regulators. Curr Opin Genet Dev. 1998     October; 8(5):539-44. -   2. Olson E. Activation of muscle-specific transcription by myogenic     helix-loop-helix proteins. Symp Soc Exp Biol. 1992; 46:331-41. -   3. Olson E N. Interplay between proliferation and differentiation     within the myogenic lineage. Dev Biol. 1992 December; 154(2):261-72. -   4. Olson E N, Perry W M. MyoD and the paradoxes of myogenesis. Curr     Biol. 1992 January; 2(1):35-7. -   5. Li L, Olson E N. Regulation of muscle cell growth and     differentiation by the -   MyoD family of helix-loop-helix proteins. Adv Cancer Res. 1992;     58:95-119. -   6. Brand T, Butler-Browne G, Fuchtbauer E M, Renkawitz-Pohl R,     Brand-Saberi B. EMBO Workshop Report: Molecular genetics of muscle     development and neuromuscular diseases Kloster Irsee, Germany, Sep.     26-Oct. 1, 1999. Embo J. 2000 May 2; 19(9):1935-41. -   7. Brand-Saberi B. Genetic and epigenetic control of skeletal muscle     development. Ann Anat. 2005 July; 187(3):199-207. -   8. Brand-Saberi B, Christ B. Genetic and epigenetic control of     muscle development in vertebrates. Cell Tissue Res. 1999 April;     296(1):199-212. -   9. Scaal M, Bonafede A, Dathe V, Sachs M, Cann G, Christ B, et al.     SF/HGF is a mediator between limb patterning and muscle development.     Development. 1999 November; 126(21):4885-93. -   10. Schwarz J J, Chakraborty T, Martin J, Zhou J M, Olson E N. The     basic region of myogenin cooperates with two transcription     activation domains to induce muscle-specific transcription. Mol Cell     Biol. 1992 January; 12(1):266-75. -   11. Christ B, Brand-Saberi B. Limb muscle development. Int J Dev     Biol. 2002; 46(7):905-14. -   12. Dutton E K, Uhm C S, Samuelsson S J, Schaffner A E, Fitzgerald S     C, Daniels M P. Acetylcholine receptor aggregation at nerve-muscle     contacts in mammalian cultures: induction by ventral spinal cord     neurons is specific to axons. J Neurosci. 1995 November;     15(11):7401-16. -   13. Daniels M P, Lowe B T, Shah S, Ma J, Samuelsson S J, Lugo B, et     al. Rodent nerve-muscle cell culture system for studies of     neuromuscular junction development: refinements and applications.     Microsc Res Tech. 2000 Apr. 1; 49(1):26-37. -   14. Uhm C S, Neuhuber B, Lowe B, Crocker V, Daniels M P.     Synapse-forming axons and recombinant agrin induce microprocess     formation on myotubes. J Neurosci. 2001 Dec. 15; 21(24):9678-89. -   15. Oakley R A, Lefcort F B, Clary D O, Reichardt L F, Prevette D,     Oppenheim R W, et al. Neurotrophin-3 promotes the differentiation of     muscle spindle afferents in the absence of peripheral targets. J     Neurosci. 1997 Jun. 1; 17(11):4262-74. -   16. Kucera J, Walro J M, Reichler J. Role of nerve and muscle     factors in the development of rat muscle spindles. Am J Anat. 1989     October; 186(2):144-60. -   17. Kucera J, Walro J. Axotomy induces fusimotor-free muscle     spindles in neonatal rats. Neurosci Lett. 1992 Mar. 2; 136(2):216-8. -   18. Albert Y, Whitehead J, Eldredge L, Carter J, Gao X, Tourtellotte     W G. Transcriptional regulation of myotube fate specification and     intrafusal muscle fiber morphogenesis. J Cell Biol. 2005 Apr. 25;     169(2):257-68. -   19. Flucher B E, Andrews S B, Daniels M P. Molecular organization of     transverse tubule/sarcoplasmic reticulum junctions during     development of excitation-contraction coupling in skeletal muscle.     Mol Biol Cell. 1994 October; 5(10):1105-18. -   20. Torgan C E, Daniels M P. Regulation of myosin heavy chain     expression during rat skeletal muscle development in vitro. Mol Biol     Cell. 2001 May; 12(5):1499-508. -   21. Flucher B E, Morton M E, Froehner S C, Daniels M P. Localization     of the alpha 1 and alpha 2 subunits of the dihydropyridine receptor     and ankyrin in skeletal muscle triads. Neuron. 1990 September;     5(3):339-51. -   22. Flucher B E, Phillips J L, Powell J A, Andrews S B, Daniels M P.     Coordinated development of myofibrils, sarcoplasmic reticulum and     transverse tubules in normal and dysgenic mouse skeletal muscle, in     vivo and in vitro. Dev Biol. 1992 April; 150(2):266-80. -   23. Flucher B E, Terasaki M, Chin H M, Beeler T J, Daniels M P.     Biogenesis of transverse tubules in skeletal muscle in vitro. Dev     Biol. 1991 May; 145(1):77-90. -   24. Das M, Molnar P, Gregory C, Riedel L, Jamshidi A, Hickman J J.     Long-term culture of embryonic rat cardiomyocytes on an organosilane     surface in a serum-free medium. Biomaterials. 2004 November;     25(25):5643-7. -   25. Das M, Wilson K, Molnar P, Hickman J J. Differentiation of     skeletal muscle and integration of myotubes with silicon     microstructures using serum-free medium and a synthetic silane     substrate. Nat Protoc. 2007; 2(7):1795-801. -   26. Das M, Bhargava N, Bhalkikar A, Kang J F, Hickman J J. Temporal     neurotransmitter conditioning restores the functional activity of     adult spinal cord neurons in long-term culture. Exp Neurol. 2008     January; 209(1):171-80. -   27. Das M, Bhargava N, Gregory C, Riedel L, Molnar P, Hickman J J.     Adult rat spinal cord culture on an organosilane surface in a novel     serum-free medium. In Vitro Cell Dev Biol Anim. 2005     November-December; 41(10):343-8. -   28. Das M, Gregory C A, Molnar P, Riedel L M, Wilson K, Hickman J J.     A defined system to allow skeletal muscle differentiation and     subsequent integration with silicon microstructures. Biomaterials.     2006 August; 27(24):4374-80. -   29. Das M, Molnar P, Devaraj H, Poeta M, Hickman J J.     Electrophysiological and morphological characterization of rat     embryonic motoneurons in a defined system. Biotechnol Prog. 2003     November-December; 19(6):1756-61. -   30. Das M, Patil S, Bhargava N, Kang J F, Riedel L M, Seal S, et al.     Auto-catalytic ceria nanoparticles offer neuroprotection to adult     rat spinal cord neurons. Biomaterials. 2007 April; 28(10):1918-25. -   31. Das M, Rumsey J W, Gregory C A, Bhargava N, Kang J F, Molnar P,     et al. Embryonic motoneuron-skeletal muscle co-culture in a defined     system. Neuroscience. 2007 May 11; 146(2):481-8. -   32. Wilson K, Molnar P, Hickman J. Integration of functional     myotubes with a Bio-MEMS device for non-invasive interrogation. Lab     Chip. 2007 July; 7(7):920-2. -   33. Rumsey J W, Das M, Kang J F, Wagner R, Molnar P, Hickman J J.     Tissue engineering intrafusal fibers: dose- and time-dependent     differentiation of nuclear bag fibers in a defined in vitro system     using neuregulin 1-beta-1. Biomaterials. 2008 March; 29(8):994-1004. -   34. Brewer G J, Boehler M D, Jones T T, Wheeler B C. NbActiv4 medium     improvement to Neurobasal/B27 increases neuron synapse densities and     network spike rates on multielectrode arrays. J Neurosci Methods.     2008 May 30; 170(2):181-7. -   35. Stenger D A, Georger J H, Dulcey C S, Hickman J J, Rudolph A S,     Nielsen T B, et al. Coplanar Molecular Assemblies of     Aminoalkylsilane and Perfluorinated Alkylsilane—Characterization and     Geometric Definition of Mammalian-Cell Adhesion and Growth. J Am     Chem Soc. 1992; 114(22):8435-42. -   36. Ruegg J. Calcium in muscle activation. Berlin: Springer Verlag;     1988. -   37. Franzini-Armstrong C, Protasi F. Ryanodine receptors of striated     muscles: a complex channel capable of multiple interactions. Physiol     Rev. 1997 July; 77(3):699-729. -   38. Ahern C A, Sheridan D C, Cheng W, Mortenson L, Nataraj P, Allen     P, et al. Ca2+ current and charge movements in skeletal myotubes     promoted by the beta-subunit of the dihydropyridine receptor in the     absence of ryanodine receptor type 1. Biophys J. 2003 February; 84(2     Pt 1):942-59. -   39. Sheridan D C, Carbonneau L, Ahern C A, Nataraj P, Coronado R.     Ca2+-dependent excitation-contraction coupling triggered by the     heterologous cardiac/brain DHPR beta2a-subunit in skeletal myotubes.     Biophys J. 2003 December; 85(6):3739-57. -   40. Sheridan D C, Cheng W, Ahern C A, Mortenson L, Alsammarae D,     Vallejo P, et al. Truncation of the carboxyl terminus of the     dihydropyridine receptor beta1a subunit promotes Ca2+ dependent     excitation-contraction coupling in skeletal myotubes. Biophys J.     2003 January; 84(1):220-37. -   41. Brewer G J, Torricelli J R, Evege E K, Price P J. Optimized     survival of hippocampal neurons in B27-supplemented Neurobasal, a     new serum-free medium combination. J Neurosci Res. 1993 Aug. 1;     35(5):567-76. -   42. Alterio J, Courtois Y, Robelin J, Bechet D, Martelly I. Acidic     and basic fibroblast growth factor mRNAs are expressed by skeletal     muscle satellite cells. Biochem Biophys Res Commun. 1990 Feb. 14;     166(3):1205-12. -   43. Clegg C H, Linkhart T A, Olwin B B, Hauschka S D. Growth factor     control of skeletal muscle differentiation: commitment to terminal     differentiation occurs in G1 phase and is repressed by fibroblast     growth factor. J Cell Biol. 1987 August; 105(2):949-56. -   44. Bottenstein J E. Advances in vertebrate cell culture methods.     Science. -   1988 Feb. 12; 239(4841 Pt 2):G42, G8. -   45. Bottenstein J E, Hunter S F, Seidel M. CNS neuronal cell     line-derived factors regulate gliogenesis in neonatal rat brain     cultures. J Neurosci Res. 1988 July; 20(3):291-303. -   46. Bottenstein J E. Proliferation of glioma cells in serum-free     defined medium. Cancer Treat Rep. 1981; 65 Suppl 2:67-70. -   47. Morrow N G, Kraus W E, Moore J W, Williams R S, Swain J L.     Increased expression of fibroblast growth factors in a rabbit     skeletal muscle model of exercise conditioning. J Clin Invest. 1990     June; 85(6):1816-20. -   48. Gonzalez A M, Buscaglia M, Ong M, Baird A. Distribution of basic     fibroblast growth factor in the 18-day rat fetus: localization in     the basement membranes of diverse tissues. J Cell Biol. 1990 March;     110(3):753-65. -   49. Moore J W, Dionne C, Jaye M, Swain J L. The mRNAs encoding     acidic FGF, basic FGF and FGF receptor are coordinately     downregulated during myogenic differentiation. Development. 1991     March; 111(3):741-8. -   50. Anderson J E, Liu L, Kardami E. Distinctive patterns of basic     fibroblast growth factor (bFGF) distribution in degenerating and     regenerating areas of dystrophic (mdx) striated muscles. Dev Biol.     1991 September; 147(1):96-109. -   51. Olwin B B, Rapraeger A. Repression of myogenic differentiation     by aFGF, bFGF, and K-FGF is dependent on cellular heparan sulfate. J     Cell Biol. 1992 August; 118(3):631-9. -   52. Arsic N, Zacchigna S, Zentilin L, Ramirez-Correa G, Pattarini L,     Salvi A, et al. Vascular endothelial growth factor stimulates     skeletal muscle regeneration in vivo. Mol Ther. 2004 November;     10(5):844-54. -   53. Germani A, Di Carlo A, Mangoni A, Straino S, Giacinti C, Turrini     P, et al. Vascular endothelial growth factor modulates skeletal     myoblast function. Am J Pathol. 2003 October; 163(4):1417-28. -   54. Lee E W, Michalkiewicz M, Kitlinska J, Kalezic I, Switalska H,     Yoo P, et al. Neuropeptide Y induces ischemic angiogenesis and     restores function of ischemic skeletal muscles. J Clin Invest. 2003     June; 111(12):1853-62. -   55. Lescaudron L, Peltekian E, Fontaine-Perus J, Paulin D, Zampieri     M, Garcia L, et al. Blood borne macrophages are essential for the     triggering of muscle regeneration following muscle transplant.     Neuromuscul Disord. 1999 March; 9(2):72-80. -   56. Motamed K, Blake D J, Angello J C, Allen B L, Rapraeger A C,     Hauschka S D, et al. Fibroblast growth a factor receptor-1 mediates     the inhibition of endothelial cell proliferation and the promotion     of skeletal myoblast differentiation by SPARC: a role for protein     kinase A. J Cell Biochem. 2003 Oct. 1; 90(2):408-23. -   57. Dusterhoft S, Pette D. Evidence that acidic fibroblast growth     factor promotes maturation of rat satellite-cell-derived myotubes in     vitro. Differentiation. 1999 November; 65(3):161-9. -   58. Fu X, Cuevas P, Gimenez-Gallego G, Sheng Z, Tian H. Acidic     fibroblast growth factor reduces rat skeletal muscle damage caused     by ischemia and reperfusion. Chin Med J (Engl). 1995 March;     108(3):209-14. -   59. Smith J, Schofield P N. The effects of fibroblast growth factors     in long-term primary culture of dystrophic (mdx) mouse muscle     myoblasts. Exp Cell Res. 1994 January; 210(1):86-93. -   60. Oliver L, Raulais D, Vigny M. Acidic fibroblast growth factor     (aFGF) in developing normal and dystrophic (mdx) mouse muscles.     Distribution in degenerating and regenerating mdx myofibres. Growth     Factors. 1992; 7(2):97-106. -   61. Dell'Era P, Ronca R, Coco L, Nicoli S, Metra M, Presta M.     Fibroblast growth factor receptor-1 is essential for in vitro     cardiomyocyte development. Circ Res. 2003 Sep. 5; 93(5):414-20. -   62. Husmann I, Soulet L, Gautron J, Martelly I, Barritault D. Growth     factors in skeletal muscle regeneration. Cytokine Growth Factor Rev.     1996 October; 7(3):249-58. -   63. Kurek J B, Nouri S, Kannourakis G, Murphy M, Austin L. Leukemia     inhibitory factor and interleukin-6 are produced by diseased and     regenerating skeletal muscle. Muscle Nerve. 1996 October;     19(10):1291-301. -   64. Megeney L A, Perry R L, LeCouter J E, Rudnicki M A. bFGF and LIF     signaling activates STAT3 in proliferating myoblasts. Dev Genet.     1996; 19(2):139-45. -   65. Vakakis N, Bower J, Austin L. In vitro myoblast to myotube     transformations in the presence of leukemia inhibitory factor.     Neurochem Int. 1995 October-November; 27(4-5):329-35. -   66. Martinou J C, Martinou I, Kato A C. Cholinergic differentiation     factor (CDF/LIF) promotes survival of isolated rat embryonic     motoneurons in vitro. Neuron. 1992 April; 8(4):737-44. -   67. Sun L, Ma K, Wang H, Xiao F, Gao Y, Zhang W, et al.     JAK1-STAT1-STAT3, a key pathway promoting proliferation and     preventing premature differentiation of myoblasts. J Cell Biol. 2007     Oct. 8; 179(1):129-38. -   68. Malm C, Sjodin T L, Sjoberg B, Lenkei R, Renstrom P, Lundberg I     E, et al. Leukocytes, cytokines, growth factors and hormones in     human skeletal muscle and blood after uphill or downhill running. J     Physiol. 2004 May 1; 556(Pt 3):983-1000. -   69. Zorzano A, Kaliman P, Guma A, Palacin M. Intracellular signals     involved in the effects of insulin-like growth factors and     neuregulins on myofibre formation. Cell Signal. 2003 February;     15(2):141-9. -   70. Sakuma K, Watanabe K, Sano M, Uramoto I, Totsuka T. Differential     adaptation of growth and differentiation factor 8/myostatin,     fibroblast growth factor 6 and leukemia inhibitory factor in     overloaded, regenerating and denervated rat muscles. Biochim Biophys     Acta. 2000 Jun. 2; 1497(1):77-88. -   71. Biesecker G. The complement SC5b-9 complex mediates cell     adhesion through a vitronectin receptor. J Immunol. 1990 Jul. 1;     145(1):209-14. -   72. Gullberg D, Sjoberg G, Veiling T, Sejersen T. Analysis of     fibronectin and vitronectin receptors on human fetal skeletal muscle     cells upon differentiation. Exp Cell Res. 1995 September;     220(1):112-23. -   73. Wang X, Wu H, Zhang Z, Liu S, Yang J, Chen X, et al. Effects of     interleukin-6, leukemia inhibitory factor, and ciliary neurotrophic     factor on the proliferation and differentiation of adult human     myoblasts. Cell Mol Neurobiol. 2008 January; 28(1):113-24. -   74. Chen X, Mao Z, Liu S, Liu H, Wang X, Wu H, et al.     Dedifferentiation of adult human myoblasts induced by ciliary     neurotrophic factor in vitro. Mol Biol Cell. 2005 July;     16(7):3140-51. -   75. Chen X P, Liu H, Liu S H, Wu Y, Wu H T, Fan M. [Exogenous rhCNTF     inhibits myoblast differentiation of skeletal muscle of adult human     in vitro]. Sheng Li Xue Bao. 2003 Aug. 25; 55(4):464-8. -   76. Cannon J G. Intrinsic and extrinsic factors in muscle aging. Ann     N Y Acad Sci. 1998 Nov. 20; 854:72-7. -   77. Marques M J, Neto H S. Ciliary neurotrophic factor stimulates in     vivo myotube formation in mice. Neurosci Lett. 1997 Sep. 26;     234(1):43-6. -   78. Carrasco D I, English A W. Neurotrophin 4/5 is required for the     normal development of the slow muscle fiber phenotype in the rat     soleus. J Exp Biol. 2003 July; 206(Pt 13):2191-200. -   79. Simon M, Porter R, Brown R, Coulton G R, Terenghi G. Effect of     NT-4 and BDNF delivery to damaged sciatic nerves on phenotypic     recovery of fast and slow muscles fibres. Eur J Neurosci. 2003     November; 18(9):2460-6. -   80. Choi-Lundberg D L, Bohn M C. Ontogeny and distribution of glial     cell line-derived neurotrophic factor (GDNF) mRNA in rat. Brain Res     Dev Brain Res. 1995 Mar, 16; 85(1):80-8. -   81. Lin L F, Doherty D H, Lile J D, Bektesh S, Collins F. GDNF: a     glial cell line-derived neurotrophic factor for midbrain     dopaminergic neurons. Science. 1993 May 21; 260(5111):1130-2. -   82. Yang L X, Nelson P G. Glia cell line-derived neurotrophic factor     regulates the distribution of acetylcholine receptors in mouse     primary skeletal muscle cells. Neuroscience. 2004; 128(3):497-509. -   83. Golden J P, DeMaro J A, Osborne P A, Milbrandt J, Johnson E M,     Jr. Expression of neurturin, GDNF, and GDNF family-receptor mRNA in     the developing and mature mouse. Exp Neurol. 1999 August;     158(2):504-28. -   84. Henderson C E, Phillips H S, Pollock R A, Davies A M, Lemeulle     C, Armanini M, et al. GDNF: a potent survival factor for motoneurons     present in peripheral nerve and muscle. Science. 1994 Nov. 11;     266(5187):1062-4. -   85. Heinrich G. A novel BDNF gene promoter directs expression to     skeletal muscle. BMC Neurosci. 2003 Jun. 18; 4:11. -   86. Mousavi K, Parry D J, Jasmin B J. BDNF rescues myosin heavy     chain IIB muscle fibers after neonatal nerve injury. Am J Physiol     Cell Physiol. 2004 July; 287(1):C22-9. -   87. Chen J, von Bartheld C S. Role of exogenous and endogenous     trophic factors in the regulation of extraocular muscle strength     during development. Invest Ophthalmol Vis Sci. 2004 October;     45(10):3538-45. -   88. Bordet T, Lesbordes J C, Rouhani S, Castelnau-Ptakhine L,     Schmalbruch H, Haase G, et al. Protective effects of cardiotrophin-1     adenoviral gene transfer on neuromuscular degeneration in transgenic     ALS mice. Hum Mol Genet. 2001 Sep. 1; 10(18):1925-33. -   89. Dolcet X, Soler R M, Gould T W, Egea J, Oppenheim R W, Comella     J X. Cytokines promote motoneuron survival through the Janus     kinase-dependent activation of the phosphatidylinositol 3-kinase     pathway. Mol Cell Neurosci. 2001 December; 18(6):619-31. -   90. Lesbordes J C, Bordet T, Haase G, Castelnau-Ptakhine L, Rouhani     S, Gilgenkrantz H, et al. In vivo electrotransfer of the     cardiotrophin-1 gene into skeletal muscle slows down progression of     motor neuron degeneration in pmn mice. Hum Mol Genet. 2002 Jul. 1;     11(14):1615-25. -   91. Nishikawa J, Sakuma K, Sorimachi Y, Yoshimoto K, Yasuhara M.     Increase of Cardiotrophin-1 immunoreactivity in regenerating and     overloaded but not denervated muscles of rats. Neuropathology. 2005     March; 25(1):54-65. -   92. Mitsumoto H, Klinkosz B, Pioro E P, Tsuzaka K, Ishiyama T,     O'Leary R M, et al. Effects of cardiotrophin-1 (CT-1) in a mouse     motor neuron disease. Muscle Nerve. 2001 June; 24(6):769-77. -   93. Oppenheim R W, Wiese S, Prevette D, Armanini M, Wang S, Houenou     L J, et al. Cardiotrophin-1, a muscle-derived cytokine, is required     for the survival of subpopulations of developing motoneurons. J     Neurosci. 2001 Feb. 15; 21(4):1283-91. -   94. Peroulakis M E, Forger N G. Ciliary neurotrophic factor     increases muscle fiber number in the developing levator ani muscle     of female rats. Neurosci Lett. -   2000 Dec. 22; 296(2-3):73-6. -   95. Sheng Z, Pennica D, Wood W I, Chien K R. Cardiotrophin-1     displays early expression in the murine heart tube and promotes     cardiac myocyte survival. Development. 1996 February; 122(2):419-28. -   96. Jaworska-Wilczynska M, Wilczynski G M, Engel W K, Strickland D     K, Weisgraber K H, Askanas V. Three lipoprotein receptors and     cholesterol in inclusion-body myositis muscle. Neurology. 2002 Feb.     12; 58(3):438-45. -   97. Caratsch C G, Santoni A, Eusebi F. Interferon-alpha, beta and     tumor necrosis factor-alpha enhance the frequency of miniature     end-plate potentials at rat neuromuscular junction. Neurosci Lett.     1994 Jan. 17; 166(1):97-100. -   98. Al-Shanti N, Saini A, Faulkner S H, Stewart C E. Beneficial     synergistic interactions of TNF-alpha and IL-6 in C2 skeletal     myoblasts-potential cross-talk with IGF system. Growth Factors. 2008     April; 26(2):61-73. -   99. Fowler V M, Sussmann M A, Miller P G, Flucher B E, Daniels M P.     Tropomodulin is associated with the free (pointed) ends of the thin     filaments in rat skeletal muscle. J Cell Biol. 1993 January;     120(2):411-20. -   100. Jin P, Sejersen T, Ringertz N R. Recombinant platelet-derived     growth factor-BB stimulates growth and inhibits differentiation of     rat L6 myoblasts. J Biol Chem. 1991 Jan. 15; 266(2):1245-9. -   101. Kudla A J, John M L, Bowen-Pope D F, Rainish B, Olwin B B. A     requirement for fibroblast growth factor in regulation of skeletal     muscle growth and differentiation cannot be replaced by activation     of platelet-derived growth factor signaling pathways. Mol Cell Biol.     1995 June; 15(6):3238-46. -   102. Quinn L S, Ong L D, Roeder R A. Paracrine control of myoblast     proliferation and differentiation by fibroblasts. Dev Biol. 1990     July; 140(1):8-19. -   103. Yablonka-Reuveni Z. Development and postnatal regulation of     adult myoblasts. Microsc Res Tech. 1995 Apr. 1; 30(5):366-80. -   104. Gold M R. The effects of vasoactive intestinal peptide on     neuromuscular transmission in the frog. J Physiol. 1982 June;     327:325-35. -   105. Gozes I, Steingart R A, Spier A D. NAP mechanisms of     neuroprotection. J Mol Neurosci. 2004; 24(1):67-72. -   106. Aracil A, Belmonte C, Calo G, Gallar J, Gozes I, Hoyer D, et     al. Proceedings of Neuropeptides 2004, the XIV European     Neuropeptides Club meeting. Neuropeptides. 2004 December;     38(6):369-71. -   107. Robertson T A, Dutton N S, Martins R N, Taddei K, Papadimitriou     J M. Comparison of astrocytic and myocytic metabolic dysregulation     in apolipoprotein E deficient and human apolipoprotein E transgenic     mice. Neuroscience. 2000; 98(2):353-9. -   108. Langen R C, Schols A M, Kelders M C, Wouters E F,     Janssen-Heininger Y M. Enhanced myogenic differentiation by     extracellular matrix is regulated at the early stages of myogenesis.     In Vitro Cell Dev Biol Anim. 2003 March-April; 39(3-4):163-9. -   109. Foster R F, Thompson J M, Kaufman S J. A laminin substrate     promotes myogenesis in rat skeletal muscle cultures: analysis of     replication and development using antidesmin and anti-BrdUrd     monoclonal antibodies. Dev Biol. 1987 July; 122(1):11-20. -   110. Hantai D, Rao J S, Reddy B R, Festoff B W. Developmental     appearance of thrombospondin in neonatal mouse skeletal muscle. Eur     J Cell Biol. 1991 August; 55(2):286-94. -   111. Kuhl U, Ocalan M, Timpl R, von der Mark K. Role of laminin and     fibronectin in selecting myogenic versus fibrogenic cells from     skeletal muscle cells in vitro. Dev Biol. 1986 October;     117(2):628-35. -   112. Lyles J M, Amin W, Weill C L. Matrigel enhances myotube     development in a serum-free defined medium. Int J Dev Neurosci.     1992; 10(1):59-73. -   113. Song W K, Wang W, Foster R F, Bielser D A, Kaufman S J.     H36-alpha 7 is a novel integrin alpha chain that is developmentally     regulated during skeletal myogenesis. J Cell Biol. 1992 May;     117(3):643-57. -   114. Swasdison S, Mayne R. Formation of highly organized skeletal     muscle fibers in vitro. Comparison with muscle development in vivo.     J Cell Sci. 1992 July; 102 (Pt 3):643-52. -   115. Wang P, Yang G, Mosier D R, Chang P, Zaidi T, Gong Y D, et al.     Defective neuromuscular synapses in mice lacking amyloid precursor     protein (APP) and APP-Like protein 2. J Neurosci. 2005 Feb. 2;     25(5):1219-25. -   116. Yang L, Wang B, Long C, Wu G, Zheng H. Increased asynchronous     release and aberrant calcium channel activation in amyloid precursor     protein deficient neuromuscular synapses. Neuroscience. 2007 Nov.     23; 149(4):768-78. -   117. Akaaboune M, Allinquant B, Farza H, Roy K, Magoul R, Fiszman M,     et al. Developmental regulation of amyloid precursor protein at the     neuromuscular junction in mouse skeletal muscle. Mol Cell Neurosci.     2000 April; 15(4):355-67. -   118. Hall B K, Miyake T. All for one and one for all: condensations     and the initiation of skeletal development. Bioessays. 2000     February; 22(2):138-47. -   119. Fan C M, Tessier-Lavigne M. Patterning of mammalian somites by     surface ectoderm and notochord: evidence for sclerotome induction by     a hedgehog homolog. Cell. 1994 Dec. 30; 79(7):1175-86. -   120. Munsterberg A E, Kitajewski J, Bumcrot D A, McMahon A P, Lassar     A B. Combinatorial signaling by Sonic hedgehog and Wnt family     members induces myogenic bHLH gene expression in the somite. Genes     Dev. 1995 Dec. 1; 9(23):2911-22. -   121. Nelson C E, Morgan B A, Burke A C, Laufer E, DiMambro E,     Murtaugh L C, et al. Analysis of Hox gene expression in the chick     limb bud. Development. 1996 May; 122(5):1449-66. -   122. Cossu G, Tajbakhsh S, Buckingham M. How is myogenesis initiated     in the embryo? Trends Genet. 1996 June; 12(6):218-23. -   123. Currie P D, Ingham P W. Induction of a specific muscle cell     type by a hedgehog-like protein in zebrafish. Nature. 1996 Aug. 1;     382(6590):452-5. -   124. Norris W, Neyt C, Ingham P W, Currie P D. Slow muscle induction     by Hedgehog signalling in vitro. J Cell Sci. 2000 August; 113 (Pt     15):2695-703. -   125. Elia D, Madhala D, Ardon E, Reshef R, Halevy 0. Sonic hedgehog     promotes proliferation and differentiation of adult muscle cells:     Involvement of MAPK/ERK and PI3K/Akt pathways. Biochim Biophys Acta.     2007 September; 1773(9): 1438-46. -   126. Pagan S M, Ros M A, Tabin C, Fallon J F. Surgical removal of     limb bud Sonic hedgehog results in posterior skeletal defects. Dev     Biol. 1996 Nov. 25; 180(1):35-40. -   127. Holler, F. James; Skoog, Douglas A; Crouch, Stanley R (2007).     “Chapter 1”. Principles of Instrumental Analysis (6th Edition ed.).     Cengage Learning. p. 9. ISBN 9780495012016. -   128. King T, Pozzi M, Manara A (2000). Piezoactuators for     ‘real-world’ applications—Can they deliver sufficient displacement?     Power Engineering J, 14, 3: 105-110 -   129. X. J. Lou (2009), Polarization fatigue in ferroelectric thin     films and related materials. 105, 024101-1 -   130. Madhu Santosh Ku Mutyala•Deepika Bandhanadham•Liu Pan•Vijaya     Rohini Pendyala•Hai-Feng Ji ( ). Mechanical and electronic     approaches to improve the sensitivity of microcantilever sensors.     Acta Mech Sin, 25:1-12 -   131. Das, Kerry Wilson, Peter Molnar and James J Hickman ( ).     Differentiation of skeletal muscle and integration of myotubes with     silicon microstructures using serum-free medium and a synthetic     silane substrate. Nature Protocols 2.7, 1795(7). -   132. Philip S. Waggoner and Harold G. Craighead (2007). Micro- and     nanomechanical sensors for environmental, chemical, and biological     detection. Lab Chip, 7, 1238-1255 -   133. Bren-Mattison Y, Olwin B B. Sonic hedgehog inhibits the     terminal differentiation of limb myoblasts committed to the slow     muscle lineage. Dev Biol. 2002 Feb. 15; 242(2):130-48. -   134. Maves L, Waskiewicz A J, Paul B, Cao Y, Tyler A, Moens C B, et     al. Pbx homeodomain proteins direct Myod activity to promote     fast-muscle differentiation. Development. 2007 September;     134(18):3371-82. -   135. Koleva M, Kappler R, Vogler M, Herwig A, Fulda S, Hahn H.     Pleiotropic effects of sonic hedgehog on muscle satellite cells.     Cell Mol Life Sci. 2005 August; 62(16):1863-70. -   136. Bandi E, Jevsek M, Mars T, Jurdana M, Formaggio E,     Sciancalepore M, et al. Neural agrin controls maturation of the     excitation-contraction coupling mechanism in human myotubes     developing in vitro. Am J Physiol Cell Physiol. 2008 January;     294(1):C66-73. -   137. Sanes J R. Genetic analysis of postsynaptic differentiation at     the vertebrate neuromuscular junction. Curr Opin Neurobiol. 1997     February; 7(1):93-100.

TABLE 1 Medium Composition 1 S. Cata- Refer- No Component Amount logue # Source ences 1. Neurobasal 500 ml 10888 Gibco/ [41] Invitrogen 2. Antibiotic- 5 ml 15240-062 Gibco/ Antimycotic Invitrogen 3. G5 Supplement 5 ml 17503-012 Gibco/ [42-51] (100X) Invitrogen 4. VEGF_(165 r Human) 10 μg P2654 Gibco/ [52-55] Invitrogen 5. Acidic FGF 12.5 μg 13241-013 Gibco/ [42, 49, Invitrogen 51, 56-61] 6. Heparin Sulfate 50 μg D9809 Sigma [42, 49, 51, 56-61] 7. LIF 10 μg L5158 Sigma [62-70] 8. Vitronectin 50 μg V0132 Sigma [71, 72] (Rat Plasma) 9. CNTF 20 μg CRC 401B Cell [73-77] Sciences 10. NT-3 10 μg CRN 500B Cell [15] Sciences 11. NT-4 10 μg CRN 501B Cell [78, 79] Sciences 12. GDNF 10 μg CRG 400B Cell [80-84] Sciences 13. BDNF 10 μg CRB 600B Cell [79, 85, Sciences 86] 14. CT-1 10 μg CRC 700B Cell [87-95] Sciences

TABLE 2 Medium Composition 2 Refer- No Component(s) Amount Catalog Source ences 1 Neurobasal 500 ml 10888 Invitrogen/ [41] Gibco 2 Antibiotic- 5 ml 15240- Invitrogen/ antimycotic 062 Gibco 3 Cholesterol 5 ml 12531 Invitrogen/ [96] (250X) Gibco 4 TNF-alpha, 10 μg T6674 Sigma- [97-99] human Aldrich 5 PDGF BB 50 μg P4056 Sigma- [62, 100- Aldrich 103] 6 Vasoactive 250 μg V6130 Sigma- [104] intestinal Aldrich peptide (VIP) 7 Insulin-like 25 μg I2656 Sigma- [68, 69, 98] growth Aldrich factor 1 8 NAP 1 mg 61170 AnaSpec, [105, 106] Inc. 9 r- 50 μg P2002 Panvera, [107] Apolipoprotein Madison, WI E2 10 Laminin, 2 mg 08-125 Millipore [108-114] mouse purified 11 Beta amyloid 1 mg AG966 Millipore [115-117] (1-40) 12 Human 100 μg CC065 Millipore [118] Tenascin- C protein 13 rr-Sonic 50 μg 1314-SH R&D [7, 119-129] hedgehog, Systems Shh N-terminal 14 rr-Agrin 50 μg 550-AG- R&D [130, 131] (C terminal) 100 Systems 

1-22. (canceled)
 23. A method of maintaining a muscle cell culture, the method comprising: maintaining the muscle cell culture in a serum-free medium comprising NBActiv4.
 24. The method of claim 23, wherein the muscle cell culture is maintained in a serum-free medium comprising NBActiv4 for at least 30 days.
 25. The method of claim 24, wherein the muscle cell culture is maintained in a serum-free medium comprising NBActiv4 for at least 50 days.
 26. The method of claim 23, further comprising plating the muscle cell culture onto a non-biological growth substrate prior to maintaining the muscle cell culture in a serum-free medium comprising NBActiv4.
 27. The method of claim 26, wherein the non-biological growth substrate comprises a silane molecule.
 28. The method of claim 27, wherein the non-biological growth substrate is DETA.
 29. The method of claim 23, further comprising plating the muscle cell culture at a density of 700 to 1000 muscle cells per square millimeter prior to maintaining the muscle cell culture in a serum-free medium comprising NBActiv4.
 30. The method of claim 23, further comprising replenishing the NBActiv4 every 1 to 5 days.
 31. The method of claim 30, further comprising replenishing the NBActiv4 every 3 days.
 32. The method of claim 23, further comprising maintaining the muscle cell culture in a first serum-free medium lacking NBActiv4 prior to maintaining the muscle cell culture in a second serum-free medium comprising NBActiv4.
 33. The method of claim 32, wherein the first serum-free medium comprises a factor selected from the group consisting of VEGF, acidic FGF, heparin sulfate, LIF, vitronectin, CNTF, NT-3, NT-4, GDNF, BDNF, CT-1, cholesterol, TNF-alpha, BDGF, vasoactive intestinal peptide, insulin-like growth factor 1, NAP, r-Apolipoprotein, laminin, beta amyloid, tenascin-C protein, rr-sonic hedgehog, and rr-agrin.
 34. The method of claim 33, wherein the first serum-free medium comprises vitronectin.
 35. The method of claim 34, wherein the first serum-free medium is a mixture of approximately equal volumes of the medium composition of Table 1 and the medium composition of Table
 2. 36. The method of claim 32, wherein the muscle cell culture is maintained in a first serum-free medium lacking NBActiv4 from 2 to 6 days.
 37. The method of claim 36, wherein the muscle cell culture is maintained in a first serum-free medium lacking NBActiv4 for 4 days.
 38. A cell culture system comprising; a non-biological growth substrate comprising a silane molecule; a muscle cell culture in contact with the non-biological growth substrate; and a serum-free medium comprising NBActiv4.
 39. The cell culture system of claim 38, wherein the non-biological growth substrate is DETA.
 40. The cell culture system of claim 38, wherein the muscle cell culture is at least 25% positive for neonatal MHC.
 41. The cell culture system of claim 38, wherein the muscle cell culture is at least 30 days old.
 42. The cell culture system of claim 41, wherein the muscle cell culture spontaneously contracts. 